University of Tennessee, Knoxville University of Tennessee, Knoxville TRACE: Tennessee Research and Creative TRACE: Tennessee Research and Creative Exchange Exchange Microbiology Publications and Other Works Microbiology 5-17-2019 Urea Is Both a Carbon and Nitrogen Source for Microcystis Urea Is Both a Carbon and Nitrogen Source for Microcystis aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions Lauren E. Krausfeldt University of Tennessee, Knoxville Abigail T. Farmer University of Tennessee, Knoxville Hector F. Castro Gonzlez University of Tennessee, Knoxville Brittany N. Zerpernick University of Tennessee, Knoxville Shawn R. Campagna University of Tennessee, Knoxville See next page for additional authors Follow this and additional works at: https://trace.tennessee.edu/utk_micrpubs Recommended Citation Recommended Citation Krausfeldt LE, Farmer AT, Castro Gonzalez HF, Zepernick BN, Campagna SR and Wilhelm SW (2019) Urea Is Both a Carbon and Nitrogen Source for Microcystis aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions. Frontiers in Microbiology 10:1064. doi: 10.3389/fmicb.2019.01064 This Article is brought to you for free and open access by the Microbiology at TRACE: Tennessee Research and Creative Exchange. It has been accepted for inclusion in Microbiology Publications and Other Works by an authorized administrator of TRACE: Tennessee Research and Creative Exchange. For more information, please contact [email protected].
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University of Tennessee, Knoxville University of Tennessee, Knoxville
TRACE: Tennessee Research and Creative TRACE: Tennessee Research and Creative
Exchange Exchange
Microbiology Publications and Other Works Microbiology
5-17-2019
Urea Is Both a Carbon and Nitrogen Source for Microcystis Urea Is Both a Carbon and Nitrogen Source for Microcystis
aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions
Lauren E. Krausfeldt University of Tennessee, Knoxville
Abigail T. Farmer University of Tennessee, Knoxville
Hector F. Castro Gonzlez University of Tennessee, Knoxville
Brittany N. Zerpernick University of Tennessee, Knoxville
Shawn R. Campagna University of Tennessee, Knoxville
See next page for additional authors
Follow this and additional works at: https://trace.tennessee.edu/utk_micrpubs
Recommended Citation Recommended Citation Krausfeldt LE, Farmer AT, Castro Gonzalez HF, Zepernick BN, Campagna SR and Wilhelm SW (2019) Urea Is Both a Carbon and Nitrogen Source for Microcystis aeruginosa: Tracking 13C Incorporation at Bloom pH Conditions. Frontiers in Microbiology 10:1064. doi: 10.3389/fmicb.2019.01064
This Article is brought to you for free and open access by the Microbiology at TRACE: Tennessee Research and Creative Exchange. It has been accepted for inclusion in Microbiology Publications and Other Works by an authorized administrator of TRACE: Tennessee Research and Creative Exchange. For more information, please contact [email protected].
Urea Is Both a Carbon and NitrogenSource for Microcystis aeruginosa:Tracking 13C Incorporation at BloompH ConditionsLauren E. Krausfeldt1, Abigail T. Farmer2, Hector F. Castro Gonzalez2,Brittany N. Zepernick1, Shawn R. Campagna2 and Steven W. Wilhelm1*
1 Department of Microbiology, The University of Tennessee, Knoxville, Knoxville, TN, United States, 2 Departmentof Chemistry, The University of Tennessee, Knoxville, Knoxville, TN, United States
The use of urea as a nitrogenous fertilizer has increased over the past two decades, withurea itself being readily detected at high concentrations in many lakes. Urea has beenlinked to cyanobacterial blooms as it is a readily assimilated nitrogen (N) - source forcyanobacteria that possess the enzyme urease. We tested the hypothesis that urea mayalso act as a carbon (C) source to supplemental growth requirements during the alkalineconditions created by dense cyanobacterial blooms, when concentrations of dissolvedCO2 are vanishingly low. High rates of photosynthesis markedly reduce dissolved CO2
concentrations and drive up pH. This was observed in Lake Erie during the largest bloomon record (2015) over long periods (months) and short periods (days) of time, suggestingblooms experience periods of CO2-limitation on a seasonal and daily basis. We used13C-urea to demonstrate that axenic cultures of the model toxic cyanobacterium,Microcystis aeruginosa NIES843, assimilated C at varying environmentally relevant pHconditions directly into a spectrum of metabolic pools during urea hydrolysis. Primarily,13C from urea was assimilated into central C metabolism and amino acid biosynthesispathways, including those important for the production of the hepatotoxin, microcystin,and incorporation into these pathways was at a higher percentage during growth athigher pH. This corresponded to increased growth rates on urea as the sole N sourcewith increasing pH. We propose this ability to incorporate C from urea represents yetanother competitive advantage for this cyanobacterium during dense algal blooms.
Keywords: HABs, cyanobacteria, Lake Erie, nitrogen, stable isotope probing
INTRODUCTION
Harmful cyanobacterial blooms are expanding globally across freshwater lakes and drinkingwater reservoirs. Microcystis-dominated blooms are especially prevalent and have been detectednow on six continents (Harke et al., 2016). Blooms are commonly associated with severelyimpaired water quality and the production of the class of cyanotoxins, the microcystins: secondarymetabolites originally known as “Fast Death Factor” (Bishop et al., 1959). These compoundsact as potent hepatotoxins to animals and humans, thus directly impact public health and theeconomy (Bullerjahn et al., 2016; Carmichael and Boyer, 2016). Nutrient overloading – specifically
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phosphorus (P) and nitrogen (N) – is widely accepted as aprimary cause of cyanobacterial blooms in freshwater ecosystems(EPA, 2015; Paerl et al., 2016). N inputs in particular arebecoming a major concern due the drastic global increase infertilizer consumption over the last several decades, includingthe United States (Glibert et al., 2014; Cao et al., 2017). Indeed,increased N loads have been linked to bloom formation andthe production of microcystins (Gobler et al., 2016; Paerl et al.,2016). Yet, while researchers commonly agree that nutrientconcentrations can constrain plankton biomass, less informationis available about the factors that result in specific biologicalspecies compositions.
The use of urea (CH4N2O) as a nitrogenous fertilizer hasincreased ∼100-fold in the last four decades, and now makes up>50% of the total N fertilizer consumption worldwide (Glibertet al., 2006; Paerl et al., 2016). Residual urea in soils that is notutilized by plants has historically been thought to be consumedby the terrestrial microbial community via the enzyme urease,where the byproduct ammonium can be oxidized to nitrate fordenitrification or volatilized to NH3 and lost to the atmosphere(Glibert et al., 2014). Yet various agricultural practices, such asthe application of urease inhibitors to soils, the timing of fertilizerapplication before rainfall or prior to snow melt and the broaduse of tile drainage systems, increase the likelihood that ureamay be exported to nearby aquatic systems (Glibert et al., 2006).Consequently, urea can reach concentrations as high as 150 µMand represent over 50% of the total dissolved N pool, especially insystems proximal to heavy agricultural areas (Bogard et al., 2012;Chaffin and Bridgeman, 2013; Glibert et al., 2014). Consequently,urea has been suggested to contribute to the eutrophication ofinland and coastal waters (Finlay et al., 2010; Glibert et al., 2014),and its usage has corresponded to a rise in reports of harmfulcyanobacteria in high fertilization areas (Glibert et al., 2014; Paerlet al., 2016). These observations are supported by the field studiesdemonstrating cyanobacteria can consume urea as a N source,and in some cases preferred urea over oxidized forms of N suchas nitrate (Davis et al., 2010; Chaffin and Bridgeman, 2013; Belisleet al., 2016). Studies in culture confirmed these observations andsuggest a specific role for urea in toxicity and genomic evolutionof major bloom-forming cyanobacterial taxa (Steffen et al., 2014b;Peng et al., 2018).
While the importance of urea as a N source to cyanobacteriais becoming more established, urea as a carbon (C) source hasbeen suggested but not directly examined. This source of C maybe significant during dense blooms when the dissolved CO2(aq) isdrawn down by rampant photosynthesis. CO2(aq)-limitation canmanifest during the course of a bloom, a condition observableby the proxy of increasing pH (Paerl and Huisman, 2009; Visseret al., 2016; Ji et al., 2017). This trend was clearly demonstratedin the western basin of Lake Erie during the 2015 Microcystisbloom (Figure 1). While average daily pH and phycocyaninconcentrations were variable across the summer months, bothmetrics were lower earlier in the summer (July) and graduallyincreased throughout August and September (Figures 1B,D,F).The relationship between cyanobacterial abundance and pHwas strongly and positively correlated, implying that thepH increased in parallel with cyanobacterial abundance. The
strongest correlation was observed at station WE2 located nearthe mouth of Maumee Bay (R2 = 0.87 at WE2; R2 = 0.75at WE4; R2 = 0.54 at WE8; Figures 1C,E,G), a catchmentfor the Maumee river which is responsible for the greatestagricultural nutrient loads to the western basin (Michalak et al.,2013). During this period, the average daily pH in the westernbasin ranged from ∼8.2 to 9.5, and peak phycocyanin valueswere observed at average pH values of 9.2 and higher. Theseparameters also varied over the course of the day (Figure 1H)and strongly correlated (R2 = 0.80; Figure 1I), with the highestpH values and phycocyanin concentrations consistently observedin the late afternoon.
When conditions reach such alkaline levels in lakes, theresulting inorganic C pool is primarily composed of bicarbonate(HCO3) and carbonate (CO3) (Supplementary Figure S1;Wetzel, 2001). Cyanobacteria are well equipped to handle lowCO2(aq) concentrations through the use of highly efficientC concentrating mechanisms (Price, 2011; Visser et al., 2016),although these enzymes require reducing power or ATP. Sincethe breakdown of urea by urease occurs intracellularly, the CO2from urea hydrolysis should be accessible for C-fixation and asa supplemental C source (Finlay et al., 2010; Sandrini et al.,2014). Indeed, while some may have historically thought of theCO2 produced by urea hydrolysis as a waste product (Kandeleret al., 2011), others have hypothesized that it might serve as a Csource to phototrophs (Glibert et al., 2016; Steffen et al., 2017).Urea as a C source may especially be relevant during periods ofCO2(aq)-limitation during dense blooms.
To address the above observations, we investigated the effectsof pH (as a control for inorganic C speciation) on the modeltoxic cyanobacterium M. aeruginosa during growth on urea,nitrate or ammonium as sole sources of N. Axenic cell growthwas established across a range of environmentally relevant pHconditions that mimicked progression of the bloom season.In culture experiments, 13C-labeled urea was used to traceC released during enzymatic hydrolysis and determine whetherthis cyanobacterium was capable of incorporating this C source.
MATERIALS AND METHODS
Growth of Microcystis aeruginosaCultures on Different N Sources AcrosspH ConditionsStock cultures of axenic Microcystis aeruginosa NIES843 weremaintained in 25 mL of modified CT medium (briefly, 0.05 gNa2·β-glycerophosphate, 0.04 g MgSO4·7H20, and trace metals,buffered with 0.2 g TAPS (Steffen et al., 2014b) at a pH of 8.2with 0.595 mM N (nitrate provided as KNO3/CaNO3.4H2O,ammonium provided as NH4Cl or urea) in 50 mL screw capglass culture tubes. After adjustment with NaOH to change thepH, Na+ concentrations totaled ∼1 mM at 8.2. The cultureswere incubated without shaking at 26◦C on a diel cycle (12:12 h)within the range of 50–60 µmol photon m−2 s−1 of light. Culturetubes were inverted three times daily, and caps were loosenedto allow for gas exchange. Microbial contamination checks were
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FIGURE 1 | Estimates of real-time phycocyanin concentrations (relative fluorescence units, RFU) and pH data from monitoring buoys in the western basin of LakeErie were accessed through the NOAA Great Lakes Environmental Research Laboratory website (https://www.glerl.noaa.gov/res/HABs_and_Hypoxia/rtMonSQL.php, accessed March 22, 2018). Data were collected at 15-min intervals for buoys deployed in 2015 at station WE2 and WE4 from July to September and WE8 fromAugust to September which captured before, during and after one of the biggest blooms on record in Lake Erie (Ho and Michalak, 2017). Data for phycocyanin(manufacturers reported resolution ± 0.01 RFU) and pH ( ± 0.01 units) were taken from the onboard EXO2 Multiparameter Sonde averaged over each calendar dayfor analysis to account for diel variation. Locations of each buoy used for data analysis are shown in black circles in the top left panel (A). Trends in averagephycocyanin (relative fluorescence units, RFU) and pH over the course of the summer of 2015 for WE8 (B), WE4 (D), and WE2 (F) are shown in the top middlepanels. Correlations between average daily phycocyanin and pH for WE8 (C), WE4 (E), and WE2 (G) are shown in the top right panels. Changes in pH observed at15 min intervals for the 2 weeks and Spearman correlation between pH and RFU over these 2 weeks are shown (H,I).
performed regularly by microscopy as well as for heterotrophicmicrobial growth in purity tubes of both rich and minimal media(LB and CT+ 0.05% glucose, 0.05% acetate, 0.05% pyruvate, and0.05% lactate, respectively). Total dissolved inorganic C (DIC)concentrations in the media after autoclaving, cooling andacclimation to 26◦C were measured at the Water Quality CoreFacility at the University of Tennessee using a Carbon/NitrogenAnalyzer (Shimadzau TOC-LCSH). DIC concentrations of themedium were an average of 4.3 mg/L regardless of inversion.
Prior to starting each growth curve, cells were collected on a1.0-µm nominal pore-size polycarbonate filter and resuspendedinto experimental media at a final pH of 7.7 ± 0.05, 8.2 ± 0.05,8.7 ± 0.05 or 9.2 ± 0.05 (adjusted with NaOH after autoclaving)for each respective N source. Cell concentrations were measuredusing flow cytometry (Guava easyCyteHT, Millipore) and wereused as an inoculum. Growth curves were performed at a
starting concentration of ∼75,000 cells/mL, and chlorophyll aautofluorescence (fluorescence signal units, FSU) was measuredusing a fluorometer (Turner Designs TD-700) over the courseof the experiment. Samples were examined at approximately thesame time every day for consistency after mixing by inversion.Cell concentration and FSU from preliminary growth curvesperformed the same way on different N sources correlatedstrongly and significantly (R2 = 0.96, p = 0.0001; SupplementaryFigure S2) demonstrating that FSU serves as a suitable proxyfor cell density. The additional Na+ added to achieve higherpH media conditions (at pH 9.2, there was a total of ∼1.7 mMNa+) did not have an influence on growth rate or biomassaccumulation (Supplementary Figure S3). These experimentswere performed by adding NaCl to media at a pH of 8.2so that the total Na+ concentration matched media at pHof 9.2 that was adjusted with a larger amount of NaOH. To
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determine if the cells were limited by C in the above conditions,M. aeruginosa was grown on nitrate with 0.238 mM Na2HCO3 ata pH of 8.2 and 9.2.
Tracing Cellular Metabolites and 13CIncorporation From UreaAxenic M. aeruginosa NIES 843, acclimated to growth on 12C-urea, was transferred to media containing saturated (>99%)13C-urea as the sole N source at a starting concentration of0.585 mM N in 25 mL of modified CT medium adjusted tothree different pH values using NaOH. These three values werechosen to capture a range of pH that would be observed priorto a bloom, during a bloom, and at the late stages of a bloom.For each condition, the pH was empirically determined at theonset of inoculation (after autoclaving media, which can alterpH) and resulted in pH of 7.5 ± 0.2, 8.4 ± 0.2 and 9.5 ± 0.2for experimental triplicates. The approximate proportions ofDIC in each treatment can be referenced in SupplementaryFigure S1. Cultures were inoculated at ∼25 FSU (∼75,000cells/mL) and grown for 7 days. On day 7, cell concentrationand FSU were measured (Supplementary Table S1), each culturewas filtered onto a 1.0-µm nominal pore-size polycarbonatefilter, and metabolites were immediately extracted at 4◦C withpre-chilled extraction solvent (1.3 mL of 40:40:20 HPLC gradeACN/MeOh/H2O with 0.1 M formic acid). Sample 8.4 R2 did notgrow and was excluded from the study.
Extracted metabolites were stored at −20◦C until beingdried under a stream of N2 and resuspended in sterile water.Samples were immediately placed in the Ultimate 3000 RSautosampler (Dionex, Sunnyvale, CA) and an injection volumeof 10 µL was separated through a Synergi 2.5 µm Hyrdo-RP, 100 Å, 100 × 2.00 mm liquid chromatography column(Phenomenex, Torrance, CA, United States) kept at 25◦C. Themass spectrometer was run in full scan mode following a protocoladapted from Rabinowitz (Lu et al., 2010). The chromatographiceluent was ionized via an electrospray ionization (ESI) sourcein negative mode and coupled to an Exactive Plus Orbitrapmass spectrometer (Thermo Fisher Scientific, Waltham, MA,United States) through a 0.1-mm internal diameter fused silicacapillary tube. The samples were run with a spray voltage of3 kV, N sheath gas of 10 (arbitrary units), capillary temperatureof 320◦C, and automatic gain control (AGC) target set to 3× 106
ions. Samples were analyzed at a resolution of 140,000 and ascan window of 85–800 m/z from 0 to 9 min and 110–1000 m/zfrom 9 to 25 min. Solvent A consisted of 97:3 water:methanol,10 mM tributylamine, and 15 mM acetic acid. Solvent B was 100%methanol. The solvent gradient from 0 to 5 min was 100% A 0%B, from 5 to 13 min was 80% A 20% B, from 13 to 15.5 min was45% A 55% B, from 15.5−19 min was 5% A 95% B, and from19 to 25 min was 100% A 0% B with a flow rate of 200 µL/min.Files generated by Xcalibur (RAW) were converted to theopen-source mzML format (Martens et al., 2011) via the open-source msconvert software as part of the ProteoWizard package(Chambers et al., 2012). Maven (mzroll) software, PrincetonUniversity (Melamud et al., 2010; Clasquin et al., 2012) wasused to automatically correct the total ion chromatograms based
on the retention times for each sample (Melamud et al., 2010;Clasquin et al., 2012). Metabolites were manually identified andintegrated using known masses ( ± 5 ppm mass tolerance) andretention times (1 ≤ 1.5 min). 12C- and 13C-metabolites weremanually selected and integrated by exact mass ( ± 5 ppm) andknown retention time to calculate 13C-incorporation (Lu et al.,2010). The estimated natural abundances of 13C for metaboliteswere calculated using the website https://www.sisweb.com/mstools/isotope.htm, and estimated to be within 1% of the actualvalue. Briefly, isotopic distributions (which take into accountknown natural abundances, mass and chemical formula for allisotopic variations) were used to calculate the expected percentabundance of the 13C isotope for a specific metabolite, whichwill change depending on the number of carbons. The numberswere reported as normalized values (relative abundances) to themost abundant ion for that metabolite (i.e., 12C isotope), so theserelative values had to be calculated back to absolute expectedpercentage (relative abundance of 13C divided by the total relativeabundance) manually. These were the values reported.
Data AnalysisGrowth rates of M. aeruginosa on nitrate, ammonium, andurea at different pH values were calculated in log-linear growthdetermined by log scaled FSU. Statistical differences in growth ondifferent N-forms across pH treatments were determined usinga Two-way ANOVA with post hoc pairwise comparisons usingthe Tukey Multiple Comparison test. To analyze global metabolictrends, total abundances for each metabolite were calculatedfor average technical duplicates (12C + 13C normalized by cellnumber). For NMDS analysis, abundances were log transformedand clustered using Bray Curtis similarity in Primer-e v7 (Clarkeand Gorley, 2006). Percent incorporation was determined from13C:12C ratios using all of the isotopomers and isotopologsfor each individual metabolite. Statistical differences betweentotal abundances of metabolites and percent 13C incorporationfor each metabolite were determined by One-way ANOVAfollowed by pairwise comparisons using the Tukey MultipleComparisons test.
Metatranscriptomes from Lake Erie during the 2014 Toledowater crisis (Steffen et al., 2017) at stations WE2, WE4, and WE8were evaluated for the expression of the urease alpha subunit(ureC) and carbamoyl phosphate synthase large subunit (carB).Contiguous sequences (contigs) annotated as ureC or carB bythe SEED database were downloaded along with their taxonomicclassifications from RefSeq using the MG-RAST server1. Qualitycontrolled reads (Steffen et al., 2017) from the original sampleswere recruited to the contigs, and recruited reads to eachcontig were normalized by contig length and library size toidentify proportional expression of these genes in environmentalsamples. Transcriptomes previously generated from Microcystisaeruginosa NIES843 (Steffen et al., 2014b) grown in modifiedCT media with urea, nitrate and ammonium were analyzed forexpression of C concentrating genes. Reads were downloadedfrom the National Biotechnology Information Center small readarchive (PRJNA229846). Raw reads were imported into CLC
1https://www.mg-rast.org/index.html
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Genomics Workbench and trimmed with a quality limit of 0.05,allowing for two ambiguous base pairs. Trimmed reads wererecruited to the Microcystis aeruginosa NIES843 genome at asimilarity and length fraction of 0.9 for the calculation of geneexpression (TPM).
RESULTS
Differences in the growth rates of M. aeruginosa were drivenby both the chemical species of available N and the pH of thegrowth medium (Figure 2, Supplementary Table S2). Growthrates on nitrate were not affected by pH (Figures 2A,B). Thehighest growth rate was observed on ammonium at a pH of8.2 (Figures 2C,D), but this growth rate was not sustained athigher or lower pH values. The next highest growth rate wason urea at 8.2, and this was maintained at an elevated pH of8.7 and 9.2 (Figures 2E,F). Despite differences in growth amongN treatments, the highest pH values supported the highest FSUvalues by the end of the growth curve for each N treatment, withurea and nitrate reaching an FSU greater than 600 and an averageFSU on ammonium reached greater than 400 (Figures 2A,C,E).
To determine if M. aeruginosa assimilated C from ureaduring its hydrolysis, metabolites from cells grown on 13C-ureawere examined by UPLC-HRMS. Cultures were grown at threedifferent pH values to capture the range of conditions that variedin DIC composition (7.5, 8.4, and 9.5). Global metabolite profilesdiffered between pH treatments (Supplementary Figure S4).
The greatest differences were observed in relative abundancesof metabolites between the low pH (7.5) and high pH (9.5,Figure 3, Supplementary Figure S5). In general, metabolitesnormalized by cell number were significantly lower in abundancein cells grown at high pH (Figure 3, Supplementary Figure S5).Significant fold change differences were the greatest for arginine,and significant fold changes were also observed in metabolitesimportant for arginine biosynthesis. Additionally, intermediatesin central C metabolism were significantly reduced in cellsgrown at high pH. While glutamine was not statistically differentacross treatments of varying pH, glutamate was more abundantat the lowest pH. However, significant differences in theglutamate:glutamine ratio of cultures of varying pH were notobserved (p = 0.44).
13C from urea was detected in multiple metabolites includingamino acids, their precursors and metabolites involved in centralC metabolism (Figures 4–6). Specifically, several metaboliteswith 13C signatures are key intermediates in the Calvin cycle,glycolysis and the pentose phosphate pathway (Figures 4A,C)as well as the arginine biosynthesis pathway (Figures 4B,C).In addition to arginine, other amino acids were observedto have 13C signatures, including glutamate, serine, aspartate,alanine, leucine, valine, lysine, threonine (Figures 4C, 5), andamino acid biosynthetic intermediates (Figure 6A). Notably,13C was observed in all of the precursor amino acids for theproduction of microcystins. Some 13C enriched metabolitesinvolved in oxidative stress had high percentages of incorporationsuch as glutathione (GSH), glutathione disulfide (GSSG) and
FIGURE 2 | Growth dynamics on nitrate (A), ammonium (C), and urea (E) as the sole N source at varying pH values (7.7, 8.2, 8.7, and 9.2). Growth rates arepresented below each growth curve for nitrate (B), ammonium (D), and urea (F). Stars represent = statistical significance of differences between growth rates withinthat N treatment: ∗∗∗∗p < 0.0001. All p-values generated from Two-Way ANOVA can be found in Supplementary Table S2. FSU is a measure of chlorophyll aautofluorescence (fluorescence signal units).
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FIGURE 3 | Statistically significant fold changes (p < 0.05) in abundances ofmetabolites in cells grown at high pH (9.5) compared to low pH (7.5).
ophthalmate (Figures 6B–D). 13C incorporation into GSSGwas not different across varying pH, but GSH had little orundetectable incorporation at pH of 7.5 and 8.4, and an averageincorporation of approximately 65% at pH of 9.5. 13C wasincorporated into metabolites at all pH values, but metabolitesfrom both central C metabolism and amino acid biosynthesistended to be more enriched at high pH (Figures 4–6 andSupplementary Table S3). The addition of Na+ and NaHCO3 didnot affect growth suggesting these cultures were not C starved inthese conditions.
DISCUSSION
The factors that drive proliferation and success of cyanobacteriaare numerous and poorly understood. While macronutrientenrichment (N and P) is a primary cause of blooms (Steffenet al., 2014a; Paerl et al., 2016), there are likely multiple factorscontributing to how one or more species end up dominating anecosystem. For example, the notorious Microcystis spp. persistsunder nutrient limitation by use of high-affinity transporters,production storage molecules for N and P, and use of organicsources of N and P (Harke et al., 2016). Their gas vacuoles allowfor buoyancy in the water column to influence light exposureor scavenge nutrients, and they form colonies which may helpto evade stress. They also prefer warmer water temperaturesthat are now being expanded and exacerbated by the changingclimate (Guo, 2007; Paerl and Huisman, 2008; Harke et al., 2016;Xiao et al., 2018). All of these advantages help to overcomelimitations or contribute to the use of nutrient elements, but itis ultimately C fixation or sequestration that dictates the successof phototrophs. Here, we used data generated from culture workusing the model toxic cyanobacterium M. aeruginosa NIES843,real-time pH data and publicly accessible sequence informationto demonstrate that urea can act as both a N and C source forthis cyanobacterium, an ability that may partially account for the
persistence (and perhaps even dominance) of Microcystis withinfreshwater blooms.
During algal bloom formation, an increase in pH is commonlyobserved (Paerl, 2018), as was seen in the western basin ofLake Erie during the 2015 Microcystis bloom (Figures 1B–G).Results from M. aeruginosa culture studies suggest not only thatcyanobacteria are responsible for the rise in pH, but they indeedfavor the more alkaline conditions often seen later in a bloom(Figures 1B,D,F). Cultures of M. aeruginosa, grown in bufferedmedium, did not always have higher growth rates, but the highestFSU values were seen at the higher pH values. This could indicatethat higher pH supported a higher carrying capacity, or that thesecells were more fluorescent and were in better photosynthetichealth. A preference for slightly alkaline conditions is not a newlydescribed phenomenon as cyanobacteria are typically cultured inthese conditions, but the field data and culture work presentedhere suggested that the increase in the pH during a bloom servesas a positive feedback loop: algal blooms (like those dominatedby Microcystis spp.) lead to increases in pH which lead to moreprolific cyanobacterial growth. Changes in pH during the courseof a bloom event impact DIC availability and composition,and this may have implications for species succession in lakesand other fresh waters since different Microcystis strains possessdifferent mechanisms for C concentration and prefer differentconcentrations of CO2(aq) (Sandrini et al., 2014; Sandriniet al., 2015). In addition, the success of Microcystis in alkalineconditions must be in part due to a competitive advantage againstother phytoplankton (Hutchinson, 1961). For example, diatomsthat bloom in early winter and spring in Lake Erie (Twiss et al.,2012; Hampton et al., 2017) require biogenic silica for theirfrustules (cell walls), but basic pH conditions can be corrosive tothese protective structures (Lewin, 1961; Martin–Jézéquel et al.,2000) and decrease growth rate (Lundholm et al., 2004).
Growth rates of M. aeruginosa on varying N chemistrieswere also influenced by pH (Figure 2), implying N availabilityduring the daily or seasonal alkalinity fluctuations is important.In contrast to nitrate and ammonium, elevated pH conditionson urea yielded the highest growth rates. An advantage orconvenient coincidence of growth on urea as a N source maybe that the inorganic C byproduct of urea hydrolysis can beused during times of C limitation or during conditions in whichthe DIC composition is unfavorable. However, much of theprevious work on urea-C incorporation by phytoplankton hasconcluded that urea-C is released as a waste product (Mulhollandet al., 2004; Jauzein et al., 2011); indeed the release of CO2has been historically used as a method for measuring ureaseactivity (Lund and Blackburn, 1989; Guettes et al., 2002). Whereincorporation was observed, urea-C accounted for very littleof the C uptake compared to HCO3 (Antia et al., 1977; Fanand Glibert, 2005). It is important to note that the assimilationof urea as a N source by heterotrophic bacteria in eutrophicsystems would release CO2 and could confound these results.The importance of urea-C to phototrophs may be more relevantseasonally or in varying conditions (Andersson et al., 2006), suchas the aforementioned change in the composition and decreasedavailability of inorganic C with increasing pH. The expressionof ureC during a Microcystis bloom in Lake Erie indicates
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FIGURE 4 | 13C incorporation from urea into the pathways of central carbon metabolism and pathway for arginine biosynthesis in M. aeruginosa. (A) The CalvinCycle (black arrows) with the intersecting glycolysis (gray arrows) and pentose phosphate pathways (blue arrows) adapted from White (2000) and Knoop et al.(2010). (B) The classical arginine biosynthesis pathways as adapted from Wendisch (2007). In both (A) and (B), the green filled circle indicates a metabolite in whicha 13C signature was detectable, and black filled circles indicate where the metabolite was detected but with no 13C incorporation. Open circles or no circle indicatesthis metabolite was not detectable. (C) Percent incorporation of each metabolites with detectable 13C signatures, relative to pH treatment. Red dashed linesrepresent predicted natural 13C abundance, and error bars represent one standard error. Stars represent statistical significance differences between 13Cincorporation percentages across pH treatments: ∗p < 0.01; ∗∗p < 0.05; ∗∗∗p < 0.001; ∗∗∗∗p < 0.0001. All p-values generated from One-Way ANOVA can befound in Supplementary Table S3.
that urea was primarily used by cyanobacteria rather thanheterotrophic bacteria (Supplementary Figure S6), implyingurea-derived C would be assimilated rather than released duringthis prolific bloom that shutdown the Toledo water supply(Steffen et al., 2017).
Here, it was confirmed that M. aeruginosa can directly useurea as a supplemental C source in addition to serving as a Nsource. Cultures in this experiment were grown across a rangeof environmentally relevant pHs, capturing values observedacross a typical bloom event, while still falling within the buffercapacity of the media. Notably, there were several metaboliteswith 13C signatures that are key intermediates in the Calvin cycle,pentose phosphate pathway and glycolysis, as well as amino acid
biosynthesis regardless of pH. This implies that urea as a C sourceis not only valuable when dissolved inorganic C is primarilyHCO3 and CO3; at a pH of 7.5 the inorganic C compositionwould be entirely HCO3 and CO2 and both chemistries canbe utilized by M. aeruginosa (Supplementary Figure S1). Thiswas possibly a result of the relatively low DIC concentrations inthis media compared to other common fresh water media (i.e.,BG-11) and CO2(aq) saturated or hard water lakes (Casey et al.,1998; Visser et al., 2016). However, eutrophic lakes experiencelarge fluctuations in DIC and range from CO2(aq) super-saturatedto CO2(aq) deplete (Visser et al., 2016; Morales-Williams et al.,2017). Previously published transcriptomes (Steffen et al., 2014b)from cultures grown in similar conditions at a pH of 8.2 with
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FIGURE 5 | Incorporation of 13C from urea into amino acids: serine (A), alanine (B), leucine (C), valine (D), threonine (E), and lysine (F). Red dashed lines representpredicted natural 13C abundance, and error bars represent one standard error. Stars represent statistically significant differences in 13C incorporation percentagesacross pH treatments: ∗p < 0.01. All p-values generated from One-Way ANOVA can be found in Supplementary Table S3.
FIGURE 6 | Incorporation of 13C from urea into other metabolites:3-phosphoserine (A), ophthalmate (B), glutathione (C), glutathione disulfide(D). Red dashed lines represent predicted natural 13C abundance, and errorbars represent one standard error. Stars represent statistically significantdifferences in 13C incorporation percentages across pH treatments:∗p < 0.01; ∗∗p < 0.05; ∗∗∗p < 0.001. All p-values generated from One-WayANOVA can be found in Supplementary Table S3.
the same low DIC concentrations showed that relevant CCMswere indeed active as expected: HCO3 was the dominant formof DIC at pH of 8.2 (Supplementary Figures S1, S7). Additionsof Na+, to increase highly expressed SbtA pump, or HCO3
did not improve growth rates (Supplementary Figures S2, S8),suggesting the cells were not starved for C, but were onlylikely CO2(aq)-limited (vis a vis Eldridge et al., 2004). Thisindicated the benefit of urea may be greater under conditionsin which CO2(aq) specifically is low during dense blooms thatchange system pH and therefore, alkalinity and DIC composition.Interestingly, cells grown on urea had reduced expression of sbtAcompared to cells grown on nitrate (Supplementary Figure S7),further supporting that urea is helping to satisfy C requirements.Although expression of sbtA was lower on ammonium thanboth urea and nitrate, this was likely reflective of the health ofcells which were noted to have stunted growth (Steffen et al.,2014b), but still poses interesting questions about the interactionsbetween N and C assimilation. Future studies will be neededto address the tradeoff between these two mechanisms of Cacquisition with urea as the N source. However, the greater13C incorporation percentages into several metabolites at pH 9.5compared to 7.5 and 8.4 suggests that the pH driven compositionchange in DIC in more alkaline conditions influenced theutilization of urea-C. C was likely incorporated at higher rateswith metabolites being more rapidly turned over, which issupported by higher growth rates at high pH and the generaltrend that metabolites were in lower abundances at the highpH (Figures 2, 3).
Interestingly, 13C from urea was also incorporated into theprecursor amino acids necessary for microcystin production(Figure 5). Microcystin is a heptapeptide and a N- and C-richmolecule, and the influence of urea and other N forms onmicrocystin production have been, and continue to be, studiedextensively (Davis et al., 2010; Gobler et al., 2016; Harke et al.,2016; Peng et al., 2018). Several studies provide evidence thaturea is linked to microcystin production (Finlay et al., 2010;Donald et al., 2011; Harke et al., 2016). The incorporation of 13C
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from urea into the necessary precursors for microcystin suggestsurea may serve as not only a N source for the production ofthis cyanotoxin, but also a C source. Indeed, 13C incorporationinto some of these precursor amino acids was higher at highpH; however, the microcystin concentration was below ourability for detection in this study. Again, this may be a moresignificant observation at different times of the day or seasonand is a valuable consideration to make when sampling orperforming experiments to address questions about N chemistryand microcystin production.
Another metabolite with higher 13C incorporation at pH9.5 was GSH. At a pH of 9.5, approximately 60% of the totalpool of GSH contained labeled C, whereas at the lower pHvalues 13C incorporation from urea was very little or completelyabsent. Total GSH abundance did not differ between treatments,which highlights the importance of this metabolite in Microcystisphysiology regardless of pH. Although GSH is involved inseveral metabolic processes within the cell, in phototrophs itis an important antioxidant (Smirnova and Oktyabrsky, 2005;Cameron and Pakrasi, 2010; Cassier-Chauvat and Chauvat,2014). In cyanobacteria, the ratio and concentrations ofGSH (and its oxidized form, GSSG) help to balance theredox state and buffering capacity of the cell (Smirnovaand Oktyabrsky, 2005). One of the benefits of CCMs andsubsequent increased CO2 into the carboxysome of cyanobacteriais the contribution to reducing photoinhibitory effects underlow CO2(aq) conditions by inhibiting photorespiration (Tabita,1994). A higher percentage of incorporation of 13C intoGSH may be indicative that the C from urea is valuableto cells at a higher pH by reducing the cost to maintaina cellular redox balance, thereby promoting the growth andproliferation of cells.
The model toxic cyanobacterium M. aeruginosa possesses thegenes necessary for the two mechanisms of urea assimilation(Supplementary Figure S9). The first reaction involves thecatalyzed hydrolysis of urea to NH4
+ and carbamate by theactivity of the enzyme urease. Carbamate is unstable andthought to quickly dissociate into NH4
+ and CO2 (Zimmer,2000; Mikkelsen et al., 2010). Another mechanism of ureadegradation, referred to as the elimination method, involves theuncatalyzed degradation of urea by urease and yields NH4
+ andcyanate due to the activity of the enzyme cyanase (Krajewska,2009). While this remains controversial (the elimination methodhas not been confirmed experimentally and only by in silicomodels) the end products would still be the same as hydrolyticdecomposition of urea, 2NH4
++ CO2 (Krajewska, 2009).
Although the details of this mechanism for assimilation arenot clear, incorporation of 13C from urea into intermediateswithin the Calvin cycle would suggest the point of entry ofC from urea is through C-fixation (Figure 4). Alternatively,another possible entry point of urea-C exists through activityof carbamoyl phosphate synthetase (CPS, Kim and Raushel,2004). CPS can phosphorylate carbamate, the intermediate inurea hydrolysis, to yield carbamoyl phosphate (carbamoyl-P)and this directly feeds into the arginine biosynthesis pathway,which was enriched in 13C in this study (Figure 4 andSupplementary Figure S9). M. aeruginosa NIES843 carries
genes for the large and small subunits of CPS (MAE_RS21880,CarB; MAE_RS12410, CarA). Carbamate, albeit short-lived, isalready a reduced form of C, and it would be an energeticallyfavorable mechanism of assimilation of the C from ureacompared to undergoing C fixation (which requires both ATPand reducing power). This mechanism could also serve as anentry point for N into cellular biochemistry and suggests greaterproduction of arginine, which is an important precursor forcyanophycin, a N storage molecule, and microcystin. Whilethis hypothesis needs to be experimentally confirmed, thismay have implications regarding the persistence and toxicityof Microcystis when urea is present in the environment.Indeed, carB, the large subunit of CPS, was expressed byMicrocystis during the 2014 bloom, while also expressinggenes for urea uptake and utilization (Steffen et al., 2017).Utilization of carA/carB mechanisms for C assimilation maynot be restricted to phototrophs in these systems as thesegenes are widely spread among heterotrophic bacteria as well,allowing microbes to capitalize on the urea as a readilyavailable C source via this path. However, re-examinationof Lake Erie metatranscriptomes from 2014 indicates carBexpression was primarily from cyanobacteria (SupplementaryFigure S6). In conjunction with the observation that almostall of the ureC expression was from cyanobacteria, thissuggests that the mechanism of entry for C would beenvironmentally relevant.
Regardless of the mechanism for urea-C assimilation, growingon urea may be cost effective to a cell under C-limitation,as inorganic C is “freely” given. Assuming a “Redfield ratio”of C:N of 7:1 as a conserved quota for Microcystis cells(Redfield, 1958), growth on urea could support 7.6% of acell’s C requirements. Thus, while not supplying the total Cneeds of a cell, urea may be particularly important duringperiods of the day where conditions are prime for growth(sufficient light, higher temperatures) yet dissolved CO2 hasbecome vanishingly low due to physiochemical conditions (i.e.,elevated pH) and competition. Indeed, the CCMs are active inlow DIC conditions when growing on urea (SupplementaryFigure S7) suggesting that urea-C would be supplementing theC requirement of the cells.
The use of urea as a N fertilizer in agriculture has increasedrapidly over the past several decades, and its role as a potentialdriver of eutrophication is important as it is commonly foundin freshwater systems (Glibert et al., 2006). In this study, wehave confirmed urea can serve as a N source to Microcystis spp.and is a preferred source of N for growth at high pH valuescommonly observed during blooms. Moreover, we show ureacan be used as a source of C for M. aeruginosa, regardless ofthe availability of other usable forms of inorganic C. It is likelythere is a greater need for C from urea at higher pH values,when dissolved CO2 in the water is absent, and this supplementalC may aid in managing the redox state and contribute to thesuccess of Microcystis for prolonged periods of time. We proposethat urea can be an energetically favorable C source that helpssupport metabolic homeostasis and photosynthetic efficiencyof Microcystis during dense blooms. While this physiologicalmechanism may not fully explain how Microcystis manages to
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Krausfeldt et al. Microcystis Assimilates Carbon From Urea
dominate planktonic communities, it does add an intriguing andrelevant piece to the puzzle.
AUTHOR CONTRIBUTIONS
LK and BZ performed the cultivation experiments. LK performedthe 13C experiment and analyzed the environmental metadata,environmental metatranscriptomes, and culture transcriptomes.LK and AF extracted the metabolites. AF, HC, and SC performedthe metabolomics. LK, SW, HC, BZ, and SC drafted themanuscript. All authors contributed to the metabolomic dataanalysis and study design.
FUNDING
This work was supported by grants from the National ScienceFoundation (DEB 1240870 to SW; IOS 1451528 and DBI 1530975
to SW and SC) and the Kenneth and Blaire Mossman Endowmentto the University of Tennessee.
ACKNOWLEDGMENTS
We thank George Bullerjahn, R. Michael McKay, Gary LeCleir,Robbie Martin, Courtney Christopher, and Cameron Thrash fordiscussions regarding this manuscript. We would also like tothank the reviewers and associate editor for the constructive andthoughtful feedback.
SUPPLEMENTARY MATERIAL
The Supplementary Material for this article can be foundonline at: https://www.frontiersin.org/articles/10.3389/fmicb.2019.01064/full#supplementary-material
REFERENCESAndersson, M. G., van Rijswijk, P., and Middelburg, J. J. (2006). Uptake of dissolved
inorganic nitrogen, urea and amino acids in the Scheldt estuary: comparisonof organic carbon and nitrogen uptake. Aquat. Microb. Ecol. 44, 303–315. doi:10.3354/ame044303
Antia, N. J., Berland, B. R., Bonin, D. J., and Maestrini, S. Y. (1977). Effects of ureaconcentration in supporting growth of certain marine microplanktonic algae.Phycologia 16, 105–111. doi: 10.2216/i0031-8884-16-1-105.1
Belisle, B. S., Steffen, M. M., Pound, H. L., Watson, S. B., DeBruyn, J. M.,Bourbonniere, R. A., et al. (2016). Urea in Lake Erie: organic nutrient sources aspotentially important drivers of phytoplankton biomass. J. Great Lakes Res. 42,599–607. doi: 10.1016/j.jglr.2016.03.002
Bishop, C., Anet, E., and Gorham, P. (1959). Isolation and identification of thefast-death factor in Microcystis aeruginosa NRC-1. Can. J. Biochem. Physiol. 37,453–471. doi: 10.1139/o59-047
Bogard, M. J., Donald, D. B., Finlay, K., and Leavitt, P. R. (2012). Distributionand regulation of urea in lakes of central North America. Freshwater Biol. 57,1277–1292. doi: 10.1111/j.1365-2427.2012.02775.x
Bullerjahn, G. S., McKay, R. M., Davis, T. W., Baker, D. B., Boyer, G. L.,D’Anglada, L. V., et al. (2016). Global solutions to regional problems: collectingglobal expertise to address the problem of harmful cyanobacterial blooms.A Lake Erie case study. Harmful Algae 54, 223–238. doi: 10.1016/j.hal.2016.01.003
Cameron, J. C., and Pakrasi, H. B. (2010). Essential role of glutathione inacclimation to environmental and redox perturbations in the cyanobacteriumSynechocystis sp. PCC 6803. Plant Physiol. 154, 1672–1685. doi: 10.1104/pp.110.162990
Cao, P., Lu, C., and Yu, Z. (2017). Historical nitrogen fertilizer use in agriculturalecosystem of the continental United States during 1850–2015: application rate,timing, and fertilizer types. Earth Syst. Sci. Data 10, 969–984. doi: 10.5194/essd-10-969-2018
Carmichael, W. W., and Boyer, G. L. (2016). Health impacts fromcyanobacteria harmful algae blooms: implications for the North AmericanGreat Lakes. Harmful Algae 54, 194–212. doi: 10.1016/j.hal.2016.02.002
Casey, G. D., Myers, D. N., and Finnegan, D. P. (1998). National water-qualityassessment of the Lake Erie-Lake St. Clair Basin, Michigan, Indiana, Ohio,Pennsylvania, and New York; environmental and hydrologic setting (No. 97-4256). Reston: US Dept. of the Interior, US Geological Survey; Branch ofInformation Services.
Cassier-Chauvat, C., and Chauvat, F. (2014). Responses to oxidative and heavymetal stresses in cyanobacteria: recent advances. Int. J. Mol. Sci. 16, 871–886.doi: 10.3390/ijms16010871
Chaffin, J. D., and Bridgeman, T. B. (2013). Organic and inorganic nitrogenutilization by nitrogen-stressed cyanobacteria during bloom conditions. J. Appl.Phycol. 26, 299–309. doi: 10.1007/s10811-013-0118-0
Chambers, M. C., Maclean, B., Burke, R., Amodei, D., Ruderman, D. L., Neumann,S., et al. (2012). A cross-platform toolkit for mass spectrometry and proteomics.Nat. Biotechnol. 30, 918–920. doi: 10.1038/nbt.2377
Clarke, K., and Gorley, R. (2006). PRIMER v6: User Manual/Tutorial (PlymouthRoutines in Multivariate Ecological Research). Plymouth: Primer-E Ltd.
Clasquin, M. F., Melamud, E., and Rabinowitz, J. D. (2012). LC-MS DataProcessing with MAVEN: a metabolomic analysis and visualization engine.Curr. Prot. Bioinform. 37, 14.11.1–14.11.23. doi: 10.1002/0471250953.bi1411s37
Davis, T. W., Harke, M. J., Marcoval, M. A., Goleski, J., Orano-Dawson,C., Berry, D. L., et al. (2010). Effects of nitrogenous compounds andphosphorus on the growth of toxic and non-toxic strains of Microcystis duringcyanobacterial blooms. Aquat. Microb. Ecol. 61, 149–162. doi: 10.3354/ame01445
Donald, D. B., Bogard, M. J., Finlay, K., and Leavitt, P. R. (2011).Comparative effects of urea, ammonium, and nitrate on phytoplanktonabundance, community composition, and toxicity in hypereutrophicfreshwaters. Limnol. Oceanogr. 56, 2161–2175. doi: 10.4319/lo.2011.56.6.2161
Eldridge, M. L., Trick, C. G., Alm, M., DiTullio, G. R., Rue, E. L., Bruland,K. W., et al. (2004). Phytoplankton community response to a manipulation ofbioavailable iron in the HNLC waters of the subtropical Pacific Ocean. Aquat.Microb. Ecol. 35, 79–91. doi: 10.3354/ame035079
EPA (2015). Preventing Eutrophication: Scientific Support for Dual NutrientCriteria. Washington, D.C: EPA.
Fan, C., and Glibert, P. M. (2005). Effects of light on nitrogen and carbon uptakeduring a Prorocentrum minimum bloom. Harmful Algae 4, 629–641. doi: 10.1016/j.hal.2004.08.012
Finlay, K., Patoine, A., Donald, D. B., Bogard, M. J., and Leavitt, P. R. (2010).Experimental evidence that pollution with urea can degrade water quality inphosphorus-rich lakes of the Northern Great Plains. Limnol. Oceanogr. 55,1213–1230. doi: 10.4319/lo.2010.55.3.1213
Glibert, P. M., Harrison, J., Heil, C., and Seitzinger, S. (2006). Escalatingworldwide use of urea–a global change contributing to coastal eutrophication.Biogeochemistry 77, 441–463. doi: 10.1007/s10533-005-3070-5
Glibert, P. M., Maranger, R., Sobota, D. J., and Bouwman, L. (2014). The HaberBosch–harmful algal bloom (HB–HAB) link. Environ. Res. Lett. 9:105001. doi:10.1088/1748-9326/9/10/105001
Glibert, P. M., Wilkerson, F. P., Dugdale, R. C., Raven, J. A., Dupont, C. L.,Leavitt, P. R., et al. (2016). Pluses and minuses of ammonium and nitrateuptake and assimilation by phytoplankton and implications for productivity
Frontiers in Microbiology | www.frontiersin.org 10 May 2019 | Volume 10 | Article 1064
Krausfeldt et al. Microcystis Assimilates Carbon From Urea
and community composition, with emphasis on nitrogen-enriched conditions.Limnol. Oceanogr. 61, 165–197. doi: 10.1002/lno.10203
Gobler, C. J., Burkholder, J. M., Davis, T. W., Harke, M. J., Johengen, T., Stow,C. A., et al. (2016). The dual role of nitrogen supply in controlling the growthand toxicity of cyanobacterial blooms. Harmful Algae 54, 87–97. doi: 10.1016/j.hal.2016.01.010
Guettes, R., Dott, W., and Eisentraeger, A. (2002). Determination of ureaseactivity in soils by carbon dioxide release for ecotoxicological evaluation ofcontaminated soils. Ecotoxicology 11, 357–364.
Guo, L. (2007). Doing battle with the green monster of Taihu Lake. Science317:1166. doi: 10.1126/science.317.5842.1166
Hampton, S. E., Galloway, A. W., Powers, S. M., Ozersky, T., Woo, K. H., Batt,R. D., et al. (2017). Ecology under lake ice. Ecol. Lett. 20, 98–111. doi: 10.1111/ele.12699
Harke, M. J., Steffen, M. M., Gobler, C. J., Otten, T. G., Wilhelm, S. W., Wood,S. A., et al. (2016). A review of the global ecology, genomics, and biogeographyof the toxic cyanobacterium, Microcystis spp. Harmful Algae 54, 4–20. doi:10.1016/j.hal.2015.12.007
Ho, J. C., and Michalak, A. M. (2017). Phytoplankton blooms in Lake Erie impactedby both long-term and springtime phosphorus loading. J. Great Lakes Res. 43,221–228. doi: 10.1016/j.jglr.2017.04.001
Hutchinson, G. E. (1961). The paradox of the plankton. Am. Nat. 95, 137–145.Jauzein, C., Collos, Y., Laabir, M., and Vaquer, A. (2011). Dark metabolism
and carbon–nitrogen uncoupling in the toxic dinoflagellate Alexandriumcatenella (Dinophyceae). Harmful Algae 11, 73–80. doi: 10.1016/j.hal.2011.08.002
Ji, X., Verspagen, J. M. H., Stomp, M., and Huisman, J. (2017). Competitionbetween cyanobacteria and green algae at low versus elevated CO2: who willwin, and why? J. Exp. Bot. 68, 3815–3828. doi: 10.1093/jxb/erx027
Kandeler, E., Poll, C., Frankenberger, W. T., and Tabatabai, M. A. (2011). Nitrogencycle enzymes. Methods Soil Enzymol. 211–245. doi: 10.2136/sssabookser9.c10
Kim, J., and Raushel, F. M. (2004). Access to the carbamate tunnel of carbamoylphosphate synthetase. Arch. Biochem. Biophys. 425, 33–41. doi: 10.1016/j.abb.2004.02.031
Knoop, H., Zilliges, Y., Lockau, W., and Steuer, R. (2010). The metabolic networkof Synechocystis sp. PCC 6803: systemic properties of autotrophic growth. Plantphysiol. 154, 410–422. doi: 10.1104/pp.110.157198
Krajewska, B. (2009). Ureases I. functional, catalytic and kinetic properties:a review. J. Mol. Catal. B Enzym. 59, 9–21. doi: 10.1016/j.molcatb.2009.01.003
Lewin, J. C. (1961). The dissolution of silica from diatom walls.Geochim. Cosmochim. Acta 21, 182–198. doi: 10.1016/S0016-7037(61)80054-9
Lu, W. Y., Clasquin, M. F., Melamud, E., Amador-Noguez, D., Caudy, A. A., andRabinowitz, J. D. (2010). Metabolomic analysis via reversed-phase ion-pairingliquid chromatography coupled to a stand alone orbitrap mass spectrometer.Anal. Chem. 82, 3212–3221. doi: 10.1021/ac902837x
Lund, B. A., and Blackburn, T. H. (1989). Urea turnover in a coastal marinesediment measured by a 14C-urea short-term incubation. J. Microbiol. Methods9, 297–308. doi: 10.1016/0167-7012(89)90072-9
Lundholm, N., Hansen, P. J., and Kotaki, Y. (2004). Effect of pH on growthand domoic acid production by potentially toxic diatoms of the generaPseudo-nitzschia and Nitzschia. Mar. Ecol. Prog. Ser. 273, 1–15. doi: 10.3354/meps273001
Martens, L., Chambers, M., Sturm, M., Kessner, D., Levander, F., Shofstahl, J., et al.(2011). mzML-a community standard for mass spectrometry data. Mol. Cell.Proteo. 10:R110.000133. doi: 10.1074/mcp.R110.000133
Martin–Jézéquel, V., Hildebrand, M., and Brzezinski, M. A. (2000). Siliconmetabolism in diatoms: implications for growth. J. Phycol. 36, 821–840. doi:10.1046/j.1529-8817.2000.00019.x
Melamud, E., Vastag, L., and Rabinowitz, J. D. (2010). Metabolomic analysis andvisualization engine for LC-MS data. Anal. Chem. 82, 9818–9826. doi: 10.1021/ac1021166
Michalak, A. M., Anderson, E. J., Beletsky, D., Boland, S., Bosch, N. S.,Bridgeman, T. B., et al. (2013). Record-setting algal bloom in Lake Eriecaused by agricultural and meteorological trends consistent with expectedfuture conditions. Proc. Natl. Acad. Sci. 110, 6448–6452. doi: 10.1073/pnas.1216006110
Mikkelsen, M., Jørgensen, M., and Krebs, F. C. (2010). The teraton challenge. Areview of fixation and transformation of carbon dioxide. Energy Environ. Sci. 3,43–81. doi: 10.1039/B912904A
Morales-Williams, A. M., Wanamaker, A. D. Jr., and Downing, J. A. (2017).Cyanobacterial carbon concentrating mechanisms facilitate sustained CO2depletion in eutrophic lakes. Biogeosciences 14, 2865–2875. doi: 10.5194/bg-14-2865-2017
Mulholland, M. R., Boneillo, G., and Minor, E. C. (2004). A comparison of Nand C uptake during brown tide (Aureococcus anophagefferens) blooms fromtwo coastal bays on the east coast of the USA. Harmful Algae 3, 361–376.doi: 10.1016/j.hal.2004.06.007
Paerl, H. W. (2018). Mitigating toxic planktonic cyanobacterial blooms in aquaticecosystems facing increasing anthropogenic and climatic pressures. Toxins10:76. doi: 10.3390/toxins10020076
Paerl, H. W., and Huisman, J. (2008). Blooms like it hot. Science 320, 57–58.doi: 10.1126/science.1155398
Paerl, H. W., and Huisman, J. (2009). Climate change: a catalyst for globalexpansion of harmful cyanobacterial blooms. Environ. Microbiol. Rep. 1, 27–37.doi: 10.1111/j.1758-2229.2008.00004.x
Paerl, H. W., Scott, J. T., McCarthy, M. J., Newell, S. E., Gardner, W.,Havens, K. E., et al. (2016). It takes two to tango: When and where dualnutrient (N & P) reductions are needed to protect lakes and downstreamecosystems. Environ. Sci. Technol. 50, 10805–10813. doi: 10.1021/acs.est.6b02575
Peng, G., Martin, R., Dearth, S., Sun, X., Boyer, G. L., Campagna, S., et al.(2018). Seasonally-relevant cool temperatures interact with N chemistry toincrease microcystins produced in lab cultures of Microcystis aeruginosaNIES-843. Environ. Sci. Technol. 52, 4127–4136. doi: 10.1021/acs.est.7b06532
Price, G. D. (2011). Inorganic carbon transporters of the cyanobacterial CO2concentrating mechanism. Photosyn. Res. 109, 47–57. doi: 10.1007/s11120-010-9608-y
Redfield, A. C. (1958). The biological control of chemical factors in theenvironment. Am. Sci. 46, 230A–221A.
Sandrini, G., Jakupovic, D., Matthijs, H. C., and Huisman, J. (2015). Strains ofthe harmful cyanobacterium Microcystis aeruginosa differ in gene expressionand activity of inorganic carbon uptake systems at elevated CO2 levels. Appl.Environ. Microbiol. 81, 7730–7739. doi: 10.1128/AEM.02295-15
Sandrini, G., Matthijs, H. C., Verspagen, J. M., Muyzer, G., and Huisman, J. (2014).Genetic diversity of inorganic carbon uptake systems causes variation in CO2response of the cyanobacterium Microcystis. ISME J. 8:589. doi: 10.1038/ismej.2013.179
Smirnova, G., and Oktyabrsky, O. (2005). Glutathione in bacteria. Biochemistry 70,1199–1211. doi: 10.1007/s10541-005-0248-3
Steffen, M. M., Belisle, B. S., Watson, S. B., Boyer, G. L., and Wilhelm, S. W. (2014a).Status, causes and controls of cyanobacterial blooms in Lake Erie. J. Great LakesRes. 40, 215–225. doi: 10.1016/j.jglr.2013.12.012
Steffen, M. M., Dearth, S. P., Dill, B. D., Li, Z., Larsen, K. M., Campagna, S. R.,et al. (2014b). Nutrients drive transcriptional changes that maintain metabolichomeostasis but alter genome architecture in Microcystis. ISME J. 8, 2080–2092.doi: 10.1038/ismej.2014.78
Steffen, M. M., Davis, T. W., McKay, R. M., Bullerjahn, G. S., Krausfeldt, L. E.,Stough, J. M., et al. (2017). Ecophysiological examination of the Lake ErieMicrocystis bloom in 2014: linkages between biology and the water supplyshutdown of Toledo. Ohio. Environ. Sci. Technol. 51, 6745–6755. doi: 10.1021/acs.est.7b00856
Tabita, F. R. (1994). “The biochemistry and molecular regulation of carbondioxide metabolism in cyanobacteria,” in The Molecular Biology ofCyanobacteria, ed. D. A. Bryant (Berlin: Springer), 437–467. doi: 10.1007/0-306-48205-3_14
Twiss, M., McKay, R., Bourbonniere, R., Bullerjahn, G., Carrick, H., Smith, R., et al.(2012). Diatoms abound in ice-covered Lake Erie: an investigation of offshorewinter limnology in Lake Erie over the period 2007 to 2010. J. Great Lakes Res.38, 18–30. doi: 10.1016/j.jglr.2011.12.008
Visser, P. M., Verspagen, J. M. H., Sandrini, G., Stal, L. J., Matthijs, H. C. P., Davis,T. W., et al. (2016). How rising CO2 and global warming may stimulate harmfulcyanobacterial blooms. Harmful Algae 54, 145–159. doi: 10.1016/j.hal.2015.12.006
Frontiers in Microbiology | www.frontiersin.org 11 May 2019 | Volume 10 | Article 1064
Krausfeldt et al. Microcystis Assimilates Carbon From Urea
Wendisch, V. F. (2007). Amino Acid Biosynthesis–Pathways, Regulation, andMetabolic Engineering, Vol. 5. Berlin: Springer.
Wetzel, R. G. (2001). Limnology: Lake and River Ecosystems. Houston: GulfProfessional Publishing.
White, D. (2000). Physiology and Biochemistry of Prokaryotes. Oxford: OxfordUniversity Press.
Xiao, M., Li, M., and Reynolds, C. S. (2018). Colony formation in thecyanobacterium Microcystis. Biol. Rev. 93, 1399–1420. doi: 10.1111/brv.12401
Zimmer, M. (2000). Molecular mechanics evaluation of the proposed mechanismsfor the degradation of urea by urease. J. Biomol. Struct. Dyn. 17, 787–797.doi: 10.1080/07391102.2000.10506568
Conflict of Interest Statement: The authors declare that the research wasconducted in the absence of any commercial or financial relationships that couldbe construed as a potential conflict of interest.
